Sterile Insect Technique (SIT) (Dyck et al., 2005; Knipling, 1955) involves mass rearing and release of sterile insects in a given area. Sterile insects mate with their wild counterparts leading to a decrease in the wild population. SIT can be used to control the population of pests. Pests are detrimental to humans or human concerns such as agriculture or livestock production. Pests are often disease vectors, or simply put, they carry and spread disease. Examples of such pests include mosquitos of the genus Aedes, principally Aedes aegypti, which can spread the dengue virus, causing dengue fever, or the yellow fever virus causing yellow fever. Anopoheles mosquitos are responsible for spreading malaria. However, despite its environmental benefits, SIT has been used successfully for only a limited number of insect pests to date.
Modern genetics can provide significant advances in current SIT programmes and may help in implementing pest management programmes that otherwise would not be possible. These advances include: a) improving the identification of released individuals, b) removing the need for radiation-sterilisation and c) providing automated sex-separation prior to release to eliminate females from the release population (“genetic sexing”) (Alphey et al., 2008; Papathanos et al., 2009).
Female lethal RIDL technology (female-specific Release of Insects carrying a Dominant Lethal, fsRIDL) is highly effective in separating sexes and has been successfully tested in laboratory, greenhouse and semi-field experiments (WO 01/39599). The present application and our work on genetic male sterility (GB2500113 and WO2013/131920) provide resourceful additions to the current RIDL system but more importantly, they effectively and sustainably address the two remaining genetic advances (a) and b) respectively) discussed above.
It is essential to be able to detect the presence of wild insects, even amongst overwhelming numbers of released sterile insects. This requires the released insects to be marked in some way, to distinguish them from wild type insects. Current marking techniques (Hagler and Jackson, 2001; Parker, 2005) mainly involve the use of coloured dyes in the larval diet that remains visible in the adults' tissues (e.g. codling moth, pink bollworm), or application of powder directly to the pupae (medfly). Although widely used, for example the Calco Red dietary dye for pink bollworm (Pectinophora gossypiella) moths (Graham and Mangum, 1971) and fluorescent powders marking the heads of adult medfly (Steiner, 1967), their application increases the cost of rearing, can increase the amount of handling required, and is prone to errors of interpretation (Hagler and Jackson, 2001; Hagler and Miller, 2002; Morrison et al., 2011; Robinson and Hendrichs, 2005).
It is also possible that a fraction of released sterile insects will lose the marker after release (Hagler and Jackson, 2001; Hagler and Miller, 2002), which would mean that on recapture they may be counted among the wild and fertile insects in the traps. Such error would not have a significant impact where large numbers of wild insects are captured, but in programmes attempting to eradicate a pest and where the wild pest is relatively infrequently captured, the presence of one such insect might provoke a costly round of quarantine and exceptional interventions. Furthermore, there are health concerns related to the effects of the powder on workers in mass-rearing facilities.
An example of a naturally occurring mutation that has been used as a genetic marker in operational SIT programmes is the white pupae (wp) mutation. It was first used in a small SIT trial in 1972-1973 against the Australian sheep blowfly (Lucille cuprina) as a sex separation mechanism (Robinson, 2002) through male-linked chromosomal translocations.
Following the successful sex separation of blowfly pupae based on pupal colour, the first medfly genetic sexing strains were constructed in 1984 by combining Y-autosome translocations developed by Robinson and van Heemert (1982) with the white pupae mutation (wp, located on chromosome 5) previously detected by Rössler (1979). However, these strains showed significant levels of genetic instability, which increased over time. As a result, some females could not be distinguished from the males and, consequently, an increasing number of females were released into the field (Franz, 2002).
Genetic markers that result in a new phenotype can be useful, but these markers are recessive and therefore apparent only when an individual carries two copies of the allele; they are, therefore, not applicable in situations where released males mate with the native wild female pest population and the progeny is being monitored. Furthermore, for monitoring purposes, the marker must be visible in adults and more importantly it must also be apparent in insects caught in traps, which might have been dead for several weeks before being examined.
Species-specific markers can be generated by isolating visible mutations in the species of interest, cloning the corresponding gene, and then rescuing the mutant phenotype by incorporating a wild-type copy of the gene through transformation. In fact the very first germ-line transformation of an insect; D. melanogaster (Rubin and Spradling, 1982) was possible due in large part to the availability of easily detectable eye colour markers that are the wild-type genes for mutated alleles affecting eye pigmentation. The first non-drosophilid transformation also took advantage of available eye colour markers. Loukeris et al. (Loukeris et al., 1995) identified Medfly transformants as phenotypic revertants of a white-eyed mutation carried by the recipient strain. However, this procedure is laborious and requires manipulation for each species separately, thus is not cost-effective. Moreover, phenotypic mutations have not been identified for all pest insects of interest, a fact that limits the potential use of an SIT approach to pest control.
Germline transformation requires a selectable marker. Fluorescent proteins have been used for this purpose in the vast majority of transgenesis work on pest insects. Expression of these proteins, under the control of a suitable regulatory sequence, provides a readily distinguishable marker for the transgenic insect. From a SIT perspective, another key feature is that such markers are in-built and heritable (Alphey et al., 2008). For some species, full sterilisation by irradiation is achieved at a dose that compromises the performance of insects (Bakri et al., 2005). In SIT programmes against these species the preferred applied dose is not fully sterilising in order to minimise this effect, resulting in some fertile or partially fertile insects being released. The fact that dye or powder markers are not heritable leads to the possibility that recaptured progeny of such ‘sterile’ insects with wild counterparts will be scored as wild.
Examples of dominant, heritable fluorescent markers conferred through transgenesis can be found in Allen et al., 2004; Allen et al., 2001; Berghammer et al., 1999; Catteruccia et al., 2005; Catteruccia et al., 2000; Fraser, 2012; Handler and Harrell, 2001; Horn et al., 2002; Koukidou et al., 2006; Morrison et al., 2011; Peloquin et al., 2000; Perera et al., 2002; Pinkerton et al., 2000; Tamura et al., 2000). Fluorescent proteins that are widely used today as transformation markers and can be subsequently used as monitoring tools for the released insects in an SIT control programme, include the jellyfish GFP (Chalfie et al., 1994; Prasher et al., 1992), variants of this gene that result in enhanced green intensities and other colours (e.g., EGFP, cyan, yellow), and the coral, Discosoma striata, red fluorescent protein (DsRed or RFP) (Matz et al., 1999).
Although variable between insects, broadly speaking, the life cycle stages of insects are egg to larva to pupa to adult. Strong expression of any fluorescent marker, at all developmental stages, but mainly at the adult stage, is highly desirable for the rationale of reliably tracking the released insects in the field. Moreover, it is desirable that expression is widespread across all body segments of an insect (the head, thorax and abdomen). Although a stably expressed fluorescent protein will naturally undergo the same degeneration process as any other protein following an insect's death in a monitoring trap, for instance, data indicates that initial stronger fluorescent expression will lead to enhanced sustainability of the fluorescence phenotype.
A generic promoter that drives expression of a fluorescent protein should lead to ubiquitous tissue expression, therefore being readily visible under the appropriate excitation filters and presumably at all developmental stages. We have used such generic promoter-enhancer sequences with success in the past; for example, ie1-hr5, from baculovirus AcNPV which gives strong expression in Ceratitis capitata (Gong et al., 2005), Anastrepha ludens (Condon et al., 2007), Bactrocera oleae (Ant et al., 2012), Aedes aegypti (Fu et al, 2007), Aedes albopictus (Labbe et al, 2012), Pectinophora gossypiella (Li et al, 2012) and Plutella xylostella (Li et al, 2012). Unambiguous marker expression in insects has been also reported using the Drosophila melanogaster polyubiquitin promoter driving expression of the GFP (Green Fluorescent Protein) in Drosophila melanogaster and Anastrepha suspensa (Handler and Harrell, 2001; Handler and Harrell, 1999). Polyubiquitin is a highly conserved gene found in most organisms and is active in all cells. However, in many insects (especially Medfly), it gives only weak or diffuse fluorescence, and if the strength was to be increased, may result in toxicity.
An artificial promoter containing three binding sites for Pax-6 homodimers in front of a TATA box (3xP3) has also been used with success as a driver for an enhanced GFP variant (EGFP) expression in the eye of the fruit fly Drosophila melanogaster and in the flour beetle Tribolium castaneum (Berghammer et al., 1999) and other insects. It expresses most strongly from the brain, eyes, and ocelli in adults but transgenic animals were also identified as larvae and pupae. However, the restricted spatial expression of this marker makes it potentially ineffective for field monitoring purposes, as trapped insects may lose significant parts of their bodies in a trap, resulting in misidentification and mis-recording of the caught insects in a control programme. Pinkerton et al. (Pinkerton et al., 2000) have reported the use of EGFP under the control of the Actin5C promoter of Drosophila melanogaster as a genetic marker for the transformation of Aedes aegypti mosquitoes. Actin5C is a cytoplasmic actin which, like polyubiquitin, is expressed in all or most cells. EGFP expression was clearly visible in embryos and larvae. Strongest expression in late-stage embryos was seen in sections of the gut, a result that was expected when using the exon 1 proximal promoter of the actin5C gene (Burn et al., 1989). Expression of EGFP was also very clear in pupae, consistent with an increase in cell division during this life stage. Expression levels in adults varied from strong throughout the entire animal to lines where fluorescence was limited to the gonads. These differences are indicative of position effects between the different lines with the expression of the transgene being reduced except in those tissues in which there is a high level of cell division. This is potentially limiting for the use of this promoter as a monitoring tool and advocates the development of a substantial number of strains.
Tamura et al. (2000) investigated the feasibility of the GAL4/UAS system in conjunction with piggyBac vector-mediated germ-line transformation for targeted gene expression. B. mori cytoplasmic actin A3 (BmA3) was used to drive the GAL4 gene, GFP was used as the reporter. The same authors showed that the expression of the GFP was much higher using the GAL4/UAS system than the GFP expression obtained with BmA3 alone.
Concerns have been raised regarding the use of such markers in the field due to the potential fitness penalties they may induce in the recipient strain (Catteruccia et al., 2003; Irvin et al., 2004).
Accordingly, it is highly desirable to use a promoter that, on the one hand, is able to drive strong expression of a fluorescent protein (at most developmental stages) while, on the other, simultaneously poses only minimal, or no, deleterious effects to the general health and well-being of the insect strain (i.e. conferring no or little fitness disadvantage). Identification of such a promoter sequence has been challenging as evidenced by the failures seen in the prior art to date. A genetic strain comprising an expression system with such a promoter will also overcome the problems associated with current methods of insect identification using dyes and powders which are hazardous to workers and also prone to misidentification.
We have now, surprisingly, discovered that an expression system comprising a promoter that, in combination with a functional protein, such as a fluorescent marker, is both tissue specific and not generally limited to a given body segment, seems to confer no apparent fitness disadvantage on insect strains transformed with the system. Moreover, the promoter overcomes the disadvantages of existing methods that use dyes and powders to monitor insects. The disadvantages overcome or ameliorated by the present system include one or more of increased cost of rearing, increased amount of handling, errors in identification due to human error or loss of marker by the insect, and health concerns related to the effects of the powders on workers in mass rearing facilities. Furthermore, as the markers are non-heritable, re-captured progeny will be counted as wild-type.